Fifty years of alkaloid biosynthesis in Phytochemistry · 2014. 8. 13. · Fifty years of alkaloid biosynthesis in Phytochemistryq Geoffrey A. Cordell⇑ Natural Products Inc., Evanston,

Phytochemistry 91 (2013) 29–51

Contents lists available at SciVerse ScienceDirect

Phytochemistry

journal homepage: www.elsevier .com/locate /phytochem

Review

Fifty years of alkaloid biosynthesis in Phytochemistry q

Geoffrey A. Cordell ⇑Natural Products Inc., Evanston, IL, USADepartment of Pharmaceutics, College of Pharmacy, University of Florida, Gainesville, FL 32610, USA

a r t i c l e i n f o

Article history:Available online 20 June 2012

Keywords:AlkaloidsBiosynthesisOverview

0031-9422/$ - see front matter � 2012 Elsevier Ltd. Ahttp://dx.doi.org/10.1016/j.phytochem.2012.05.012

q This manuscript dedicated to the Memory of Prof⇑ Address: Natural Products Inc., 9447 Hamlin Ave,

E-mail address: [email protected]

a b s t r a c t

An overview is presented of the studies related to the biosynthesis of alkaloids published in Phytochem-istry in the past 50 years.

� 2012 Elsevier Ltd. All rights reserved.

Contents

1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 30
1.1. Ornithine-derived alkaloids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 301.2. Nicotine . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 311.3. Tropane alkaloids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 331.4. Calystegines . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 341.5. Pyrrolizidine alkaloids. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 341.6. Retronecine . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 341.7. Lysine-derived alkaloids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 351.8. Phenylalanine–tyrosine-derived alkaloids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 36
1.8.1. Benzylisoquinolines and derivatives . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 36
1.9. Bisbenzylisoquinoline alkaloids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 371.10. Berberines and derivatives . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 371.11. Benzo[c]phenanthridines . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 391.12. Morphinan and related alkaloids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 401.13. Miscellaneous alkaloids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 41
1.13.1. Salsoline . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 411.13.2. Galanthamine . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 411.13.3. Onychine. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 411.13.4. Securinine . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 421.13.5. Colchicine . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 421.13.6. Erythrina alkaloids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 421.13.7. Ipecac alkaloids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 42

1.14. Tryptophan-derived alkaloids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 42
1.14.1. Simple indole alkaloids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 421.14.2. Strictosidine . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 431.14.3. Ajmaline . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 451.14.4. Vindoline and related alkaloids. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 451.14.5. Purine-derived alkaloids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 45
1.15. Miscellaneous alkaloids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 47
1.15.1. Cyclopenin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 47
ll rights reserved.

. Meinhart H. Zenk.Evanston, IL 60203, USA. Tel./fax: +1 847 674 6022.

http://dx.doi.org/10.1016/j.phytochem.2012.05.012

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30 G.A. Cordell / Phytochemistry 91 (2013) 29–51

1.15.2. Holaphyllamine . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 471.15.3. Ricinine and Hermidin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 48

2. Conclusions. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 48Acknowledgments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 48References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 48

1. Introduction

Since Phytochemistry was first published in October, 1961, it hasevolved to become a flagship international journal for publicationsin many areas of natural product chemistry and biology, includingstudies on the isolation, structure elucidation, biosynthesis, andbiological evaluation of numerous groups of natural products.One group which has received a great deal of coverage in the an-nals of Phytochemistry over the years is the alkaloids, in all of theirstructural diversity.

The first article on alkaloids was published in the first issue(Wightman et al., 1961). Subsequently, there have been severalthousand articles appearing in the journal relating to various as-pects of alkaloids, their origin, occurrence, chemistry, biosynthesis,ecology, and biology. For this contribution, the choice was made toconsider only the aspect of alkaloid biosynthesis, and to indicate,briefly, some of the publications that appeared in Phytochemistryover the past 50 years. Not all of the published papers can be cited,and not all of the alkaloids studied can be discussed. This article isnot intended, and could not be, a comprehensive overview of alka-loid biosynthesis in Phytochemistry in 50 years! Some authors havecontributed ground-breaking studies to the field of alkaloid bio-synthesis through the medium of Phytochemistry. These contribu-tions have been an integral aspect of the growth and respect thatthe journal has engendered over the years. Apologies are tenderedto the many authors whose studies on alkaloid biosynthesis werepublished in Phytochemistry, but whose work is not cited here.What is hoped is that a flavor of the coverage of alkaloid biosynthe-sis in Phytochemistry will evolve, and that some sense of the vastchanges that have occurred in the strategies and techniques beingused will be reflected. It is also hoped that there will be an appre-ciation of the crucial role that certain specific publications, and cer-tain areas of alkaloid biosynthesis research which have appeared inPhytochemistry, have made to the field over these 50 years. Theprincipal groups of alkaloids whose biosynthesis are describedare listed, and within each section the discussion is essentiallychronological. Sometimes one alkaloid serves as a representativeof a group of alkaloids, or as way to introduce how the technologiesthat have related to alkaloid biosynthesis have changed over thesepast 50 years.

The first issue of Phytochemistry contained two publications onalkaloids (Wightman et al., 1961; Fairbairn and Suwal, 1961).Incredibly, the first dealt with the biosynthesis of tryptophanand gramine in barley, Hordeum vulgare L. (Poaceae), and showedthat shikimic acid, anthranilic acid, indole acetic acid, and serinewere precursors of tryptophan, and that tryptophan was a goodprecursor of gramine (1) (Wightman et al., 1961). The second pa-per examined the metabolism of the major alkaloids in hemlock,Conium maculatum L. (Apiaceae). It demonstrated the rapidchanges in alkaloid levels which occurred within a 24 h period,and showed how the increased level of coniine (2) correspondedwith a decline in the level of c-coniceine (3) (Fairbairn andSuwal, 1961). Another paper on alkaloid biosynthesis also ap-peared in the first volume of Phytochemistry. This articleconcerned the formation of stachydrine (4), and demonstratedthat ornithine was incorporated without randomization, and thatproline was a direct precursor in mature plants of Medicagosativa L. (Fabaceae) (Essery et al., 1962).

With that meaningful start, the reports of alkaloid biosynthesisin Phytochemistry grew, and the journal became a popular venuefor the leading researchers in the field to publish some of theirmost important results on the biosynthesis of alkaloids. Twonames stand out for their numerous dynamic contributions to Phy-tochemistry, Professor Edward Leete and Professor Meinhart Zenk,both now deceased. The efforts of these groups, which resultedin numerous and diverse studies, revolutionized the thinking inseveral areas of alkaloid biosynthesis. Leete and co-workers con-ducted numerous decisive studies to clarify the tropane and nico-tine pathways, while Zenk and co-workers carried out innovativestudies at the enzyme and gene level which brought new clarity,and overturned many paradigms associated with earlier proposedbiosynthetic pathways. Zenk also provided two important over-views of alkaloids and their biosynthesis. The first of these, in1991 (Zenk, 1991), provided a perspective on the developmentsof the application of plant cell culture systems for secondarymetabolite biosynthesis in the search for the enzymes; a particularemphasis was placed on advances in elucidating the enzymes ofalkaloid biosynthesis. The second review was of the evolutionand current status of the phytochemistry of nitrogenous com-pounds (Zenk and Juenger, 2007). Based on an overview of threetypes of nitrogen-containing plant products (alkaloids, cyanogenicglucosides, and nonprotein amino acids), it was concluded that theintegration of the disciplines of chemistry, pharmacognosy, medi-cine, analytical sciences, cell biology, molecular biology, botany,and chemotaxonomy across traditional borders will lead to newachievements in phytochemistry, and presumably also inPhytochemistry.

1.1. Ornithine-derived alkaloids

An early stage in the formation of several simple alkaloid groupsis the formation of putrescine from either ornithine or arginine,and the dimerization to spermidine, from which several deriva-tives are formed, including those which are acylated; among thesealkaloids is lunarine (5). Zenk’s group (Sagner et al., 1997) showedthat lunarine (5) was formed by stereoselective phenol oxidativecoupling of N1,N10-bis(p-coumaroyl)spermidine in the seeds ofLunaria annua L. (Brassicaceae). p-Coumaric acid was formed fromL-phenylalanine via trans-cinnamic acid, and the spermidine, de-rived through putrescine, was preferentially synthesized from argi-nine. Following enzyme studies, it was suggested that acytochrome P450 might be responsible for the phenol-oxidativecoupling of N1,N10-bis(p-coumaroyl)spermidine to afford the hexa-hydrodibenzofuran ring of lunarine (5).

The first committed step in the nicotine and tropane alkaloidpathways is the formation of N-methylputrescine (6). Putres-cine:SAM N-methyltransferase (PMT) catalyzes the N-methylationof the diamine putrescine to form N-methylputrescine (6). The pmtgene of Nicotiana tabacum, under the regulation of the CaMV 35Spromoter, was introduced into the genome of a scopolamine-richDuboisia hybrid. Although N-methylputrescine levels of theresulting engineered hairy roots increased (2 to 4-fold), therewas no significant increase in alkaloid production (Moyano et al.,2002).

A phylogenetic tree of 27 PMT and 14 SPDS proteins (Biastoffet al., 2009) demonstrated the separation of plants in the

G.A. Cordell / Phytochemistry 91 (2013) 29–51 31

Solanaceae from those in the Convolvulaceae. Within the Solana-ceae, the relationships based on PMT from Nicotiana species paral-leled those from earlier chloroplast restriction site analysis. In spiteof their plant origin, the plant SPDS sequences cluster together, andare distant from the the PMT enzymes. From an evolutionary per-spective, it was considered that a pmt gene originated from spds atan early stage in the development of higher plants. The pmt genesare expressed in the roots in Nicotiana sylvestris Speq., Atropa bella-donna L., Datura stramonium L., and the calystegine-producingplants Solanum lycopersicum L. (Solanaceae) and Calystegia sepiumR.Br. (Convolvulaceae) (Biastoff et al., 2009).

1.2. Nicotine

Nicotine (7) is derived from a pyridine nucleus and an N-pyrrol-idine moiety joined at C-3 of the pyridine nucleus. The precursor-metabolite relationships of nicotine and its N0-demethyl derivative,nornicotine (8) have been well-studied over the years. Root admin-istration of [15N]nornicotine and [methyl-14C]-L-methionine to to-bacco (N. tabacum L. (Solanaceae)) plants led to incorporation intonicotine (7). The latter precursor was incorporated at a higher ratethan the former, and the relative incorporations were very similarin the root and shoot (Ladesic and Tso, 1964).

Whether nicotinic acid (pyridine-3-carboxylic acid) was com-pletely decarboxylated as an intermediate step in the process ofincorporation into nicotine (7) was the subject of speculation.Hydroponic root administration of [2,3,7-14C]nicotinic acid to N.tabacum resulted in labeled nicotine (7), which was degraded step-wise. Radioactivity was detected at positions 2 and 3 of nicotinicacid, and not at C-4, 5, and 6, showing that the pyridine ring didnot become symmetrical during nicotine biosynthesis (Scott andGlynn, 1967). Importantly, it was also demonstrated that attach-ment of the pyrrolidine moiety occurs exclusively at the carbonfrom which the carboxylic acid group is lost.

[5-14C]-DL-D1-Pyrroline-5-carboxylic acid was incorporatedsymmetrically into the pyrrolidine ring of nicotine (7) in Nicotianarustica L. (Solanaceae) after 6 h (Krampl et al. 1969). Labeling ap-peared predominantly at C-20, 47.8% and C-50 640%. However,the low level of incorporation into nicotine (7) (0.04%) suggestedthat it is probably not an intermediate on the main pathway of pyr-rolidine ring biosynthesis. The biosynthetic formation of nicotine(7) in callus tissue of N. tabacum was activated by indole acetic acid(IAA), but suppressed by 2,4-dichlorophenoxyacetic acid (2,4-D)(Furuya et al., 1971).

The biosynthesis of the pyrrolidine ring of nicotine (7) was alsostudied using short-term, steady-state exposures of N. glutinosa L.(Solanaceae) seedlings to 14CO2. The results, following degradationof the pyrrolidine ring, showed both symmetrical and unsymmet-rical labeling (Rueppel et al., 1974), leading to the suggestion of theoperation of two biosynthetic pathways, one involving a symmet-rical precursor and the other an unsymmetrical one.

A mixture of [20-3H]-(�)-nicotine (7) and [20-14C]-(±)-nicotine(7) was used to study nicotine metabolism in N. glauca Graham(Solanaceae) plants. After 3 days, 49.5% of the activity was in nor-nicotine (8), with negligible activity in anabasine (9), and a low le-vel (2%) in myosmine (10,20-dehydronornicotine) (Leete andChedekel, 1974). The radioactive nornicotine (8) comprised 48%[20-14C, 3H]-(�)-nornicotine and 52% [20-14C]-(+)-nornicotine, sug-gesting loss of H-20 if (+)-nornicotine is formed from (�)-nicotine(7). [20-14C]Myosmine was not a precursor of nornicotine (8) inN. glauca, indicating that dehydrogenation is not reversible, butmyosmine was degraded to nicotinic acid.

An early experiment using contiguously 13C-labeled precursorswas reported by Leete and Yu (1980). Examination of the 13C NMRspectra of nicotine (7) and nornicotine (8) ten days after adminis-tering N. glutinosa plants with [2,3-13C2]- and [5-14C]-DL-ornithine

showed conclusively the symmetrical incorporation of ornithineinto the pyrrolidine ring of these alkaloids. Satellites were ob-served for the C-20, 30, 40, and 50 resonances confirming the intactpresence of contiguous carbons at C-20, 30, and C-40, 50 by way ofa symmetrical intermediate.

In 1985, for the first time, the pyridine alkaloid content, repre-sented by nicotine (7), nornicotine (8), anabasine (9), and anata-bine (10), of the leaves and roots of 60 (of the 66 known)Nicotiana species was analyzed quantitatively by gas chromatogra-phy (Saitoh et al., 1985). The species were identified morphologi-cally, and all contained alkaloids. Root samples usually containedmore alkaloids than the leaves, and the alkaloid levels varied bya factor of up to 2000-fold. Nicotine (7) was dominant in the leavesof 33 of the species and in the roots of 51 species. Seeds containedtrace alkaloid levels. No correlations were observed between theproposed classification of the genus and the alkaloid content.

Adminstration of asymmetrically-labeled spermidine ([6-14C]-1,5,10-triazadecane) to N. glutinosa afforded nicotine (7) and nor-nicotine (8) in which the label was equally distributed at C-20

and C-50 of their pyrrolidine rings (Leete, 1985). Degradation ofspermidine to putrescine therefore occurs prior to incorporation.

A non-tobacco alkaloid producing cell suspension culture ofNicotiana plumbaginifolia Viv. (Solanaceae) demonstrated thecapacity to demethylate nicotine (7) to nornicotine (8) in up to53.2% yield. Light enhanced the catalytic activity, suggesting thatdemethylation is a photodependent process (Manceau et al.,1989).

DL-[5-14C]Ornithine afforded radioactive (S)-nicotine (7) andscopolamine (11) in a root culture of Duboisia leichhardtii F.Muell.(Solanaceae). Nicotine (7) was labeled equally at C-20 and C-50,and scopolamine (11) equally at C-1 and C-5 (Leete et al., 1990).This experiment supported the common biosynthetic origin forthe pyrrolidine ring of both nicotine and scopolamine (11) througha symmetrical intermediate. Chemically, an efficient method forthe conversion of scopolamine (11) to hyoscyamine (12) wasdescribed.

Leete and co-workers (Friesen et al., 1992), using root culturesof Nicotiana alata Link & Otto (Solanaceae), showed that in thepresence of high levels of nicotinic acid both alkaloid productionand the conversion of nicotine (7) to nornicotine (8) was inhibited,while anatabine (10) formation was stimulated. On the other hand,increased levels of 1-methyl-D1-pyrrolinium chloride enhancednicotine (7) production and inhibited N-demethylation. Increasingconcentrations of D1-piperideine promoted anabasine (9) and nor-nicotine (8) production.

The mechanism of nicotine N-demethylation was investigatedwith [pyrrolidine-20-14C]-nicotine (7) to produce (�)-nornicotine(8) intracellularly in up to 70% yield (Hao and Yeoman, 1996a).N-Formyl-30-nornicotine was detected as a by-product and wasnot regarded as an intermediate in nicotine N-demethylation. N-Demethylation of nicotine (7) does not involve pyrrolidine ring-opening. Subsequent studies in a cell-free tobacco system (Haoand Yeoman, 1996b) disclosed an enzyme catalyzing the biocon-version of nicotine (7) to nornicotine (8). The enzyme had a pHoptimum of 9.0–9.5, and a temperature optimum of 25–30 �C.The Vmax and the apparent Km were 7.6 � 10�2 pkat and 7.4 lMof 14C-nicotine, respectively; the enzyme may be NADPH depen-dent. The mechanism of the demethylation of nicotine (7) to nor-nicotine (8) was also studied in root cultures of N. alata(Solanaceae) using [40,40,50,50-2H4]nicotine (7) to afford labeled nor-nicotine (8) without loss of label according to 2H NMR (Botte et al.,1997). These data also supported a mechanism in which the C-50

protons of nicotine are not involved in the demethylation reaction.The N-demethylation of the piperidine homologs of nicotine

was similar, but not identical, to that of the pyrrolidine analogs.(R,S)-N-Methylanabasine (13) and (R,S)-N-methylanatabine were


effectively demethylated in a cell suspension culture of N.plumbaginifolia (Solanaceae). Both substrates accumulated in themedium, and strong stereoselectivity for the (R)-isomers was ob-served. No further degradation of the initial demethylation prod-ucts occurs (Bartholomeusz et al., 2005a). Furthermore, (R,S)-N-methylanabasine (13) does not compete with (S)-nicotine (7)demethylation.

The metabolism of nicotine (7) and N0-nornicotine (8) in N.plumbaginifolia suspension cell cultures that do not form endoge-nous nicotine was examined. Nicotine metabolism resulted in theaccumulation of N0-nornicotine (8), and six minor metabolites, fourwere identified as cotinine (20-oxo-nicotine), myosmine, N0-form-ylnornicotine, and N0-carboethoxynornicotine. Although cotininewas formed from [13C,2H3-methyl]nicotine (7) without loss of la-bel, N0-formylnornicotine was poorly (6%) labeled, and N0-carbo-ethoxynornicotine was unlabeled. [10-15N]Nornicotine affordedN0-formylnornicotine and N0-carboethoxynornicotine without lossof label. This indicates that N0-nornicotine and cotinine are deriveddirectly from nicotine (7), while N0-formylnornicotine and N0-car-boethoxynornicotine are metabolites of nornicotine (8); the latteris therefore not an intermediate in nicotine demethylation (Bartho-lomeusz et al., 2005b).

The next step after the formation of N-methylputrescine is oxi-dation of the primary amine to the aldehyde. A methylputrescineoxidase from N. tabacum L. has been cloned and characterized(Heim et al., 2007). Some progress was reported for the identifica-tion of the functionality of the genes associated with pyridine alka-loid biosynthesis in N. tabacum L. (Solanaceae) plants. The full-length cDNA clones for 34 genes were isolated and some activitiesof selected genes expressed, including one for a lysine decarboxyl-ase (Häkkinen et al., 2007).

Minimizing the content of nornicotine (8) in tobacco is a highlydesirable goal, because of the precursor relationship to the potent

laboratory carcinogen N0-nitrosonornicotine. Typically, in Nicotianaspecies, nornicotine (8) is a relatively minor alkaloid (2–5% of thetotal pyridine alkaloid pool in the mature leaf), and is probablyproduced almost entirely through the N-demethylation of nicotine,catalyzed by nicotine N-demethylase (NND). The gene CYP82E4was identified as the specific NND gene responsible, with CY-P82E5v2 as a putative minor NND gene. The discovery and charac-terization of CYP82E10, another tobacco NND gene, showed theorigin of CYP82E10 from N. sylvestris Speg. (Solanaceae) (Lewiset al., 2010). Through knockout mutations all three tobacco NNDgenes were identified. By generating a series of mutant NND geno-types, the relative contribution of each NND gene toward the nor-nicotine (8) content of the plant was assessed. Levels were muchlower than in conventional tobacco cultivars. The study suggesteda strategy for reducing the levels of this animal carcinogen intobacco.

To explore the relationships between the arginine and ornithineroutes to putrescine, RNAi methodology was used to down-regu-late ornithine decarboxylase transcript levels in transgenic N. taba-cum plants derived from an Agrobacterium rhizogenes-derived hairyroot culture system, and a disarmed Agrobacterium tumefaciens sys-tem (DeBoer et al., 2011). Significant ODC transcript down-regula-tion was observed leading to reduced nicotine (7) and increasedanatabine (10) levels in both cultured hairy roots and intact green-house-grown plants. Neither treatment with methyl jasmonate norwounding (removal of apices) of ODC-RNAi hairy roots restoredcapacity for normal nicotine synthesis in transgenic tissue,although markedly increased levels of anatabine (10) were ob-served. It was deduced that that the ODC-mediated route toputrescine plays an important role in determining the nico-tine:anatabine profile in N. tabacum, and is necessary for N. taba-cum to increase nicotine (7) levels in response to wound-associated stress.


1.3. Tropane alkaloids

Many studies on the biosynthesis of the tropane alkaloids hyo-scyamine (atropine) (12) and scopolamine (11) have been de-scribed in the past 50 years in Phytochemistry. In 1969, Cossonreported that extended periods of light stimulated scopolamine(11) production in Datura metel L. (Solanaceae) leaves (Cosson,1969). A simple and elegant, yet powerful, experiment on the bio-synthesis of hyoscyamine (12) and scopolamine (11) was the dem-onstration by Ahmad and Leete (1970) that DL-d-[N-methyl-14C,2-14C]ornithine was incorporated intact at the bridgehead carbonC-1, but that the isomer with the a-nitrogen methylated and la-beled was not.

Two early studies of tropane alkaloid production in tissue cul-ture were reported by Tabata and co-workers. Callus cultures de-rived from the roots of Scopolia parviflora (Dunn) Nakai(Solanaceae) were maintained for two years. Alkaloid productionwas much less than the intact rhizome, but could be enhancedslightly by addition of tropic acid [the precursor of the esterifyingunit of hyoscyamine (12) and scopolamine (11)] (Tabata et al.,1972). For Datura inoxia Mill. (Solanaceae) cultured on an auxin-free medium supplemented with kinetin, the regenerated rootscould be grown to complete, mostly diploid, plants. Scopolamine(11) production was observed to begin during root differenetiation,and the alkaloid pattern was restored in the regenerated plants(Hiraoka and Tabata, 1974). An investigation of the stereochemis-try of the incorporation of D- and L-hygrine labeled at the 20-posi-tion with 14C into the tropane alkaloids scopolamine (11) andatropine (12) in D. inoxia (Solanaceae) demonstrated that onlythe D-isomer was incorporated (McGaw and Woolley, 1978). Bothisomers were apparently equally incorporated into cuscohygrinein Physalis, however (McGaw and Woolley, 1979). Subsequentlyit was shown that hygrine was not a direct precursor of cuscohy-grine in Erythroxylum coca (Newquist et al. 1993). Rather, a pyrro-lidinium species with a four-carbon side chain was a directprecursor for ethyl (RS)[2,3-13C2, 3-14C]-4-(1-methyl-2-pyrrolidi-nyl)-3-oxobutanoate specifically and labeled cuscohygrine,whereas hygrine was poorly incorporated.

In order to examine the sites of tropane alkaloid biosynthesisand distribution, Zenk and co-workers (Weiler et al., 1981) devel-oped a radioimmunoassay for the detection of picomole quantitiesof scopolamine (12) using [N-methyl-3H3]-(�)-scopolamine as thetracer. Related alkaloids did not interfere, and the variability inplant and leaf distributions was reported for Datura sanguinea Ruiz.& Pav. Cultured roots of D. leichhardtii F.Muell. (Solanaceae), D.myoporoides R.Br., and D. hopwoodii F.Muell. produced both tro-pane and pyridine-type alkaloids. Root cultures of D. leichhardtiishowed the highest content of tropane alkaloids (hyoscyamine0.53%, scopolamine 1.16%) (Endo and Yamada, 1985).

An attempt was made to examine the variation in alkaloid pat-terns in the petioles of over 1000 plants of the genera Datura, Sco-polia, and Hyoscyamus, resulting in the establishment oftransformed root cultures of the high and low alkaloid-producingplants. Although relative alkaloid patterns were retained betweenspecies, less differentiation within species was observed in tissueculture, suggesting that full expression of alkaloid potential wasnot occurring (Parr et al. 1990). 5,6-Dichloro-indole acetic acid(at 0.01 mg/L) was a powerful inducer of tropane alkaloid (partic-ularly scopolamine) biosynthesis in a Duboisia hybrid root culture(Yoshimatsu et al., 1990).

Tropane alkaloid biosynthesis in hairy roots of Hyoscyamus al-bus L. (Solanaceae) transformed with A. rhizogenes was enhancedwith added KNO3 to produce up to 30 mg/100 ml of tropane alka-loids, predominantly 6b-hydroxyhyoscyamine (25%) and hyoscya-mine (12) (63%) (Sauerwein and Shimomura, 1991). High levels ofatropine esterase activity were found at all stages of development

of A. belladonna, Datura tatula L., D. leichhardtii, Hyoscyamus niger L.,and Scopolia japonica Maxim. (Kitamura et al., 1992). Evidence wasalso presented (Kitamura et al., 1996) that recycling of the tropicacid moiety of atropine (12) may be occurring in D. myoporoidesroot cultures.

In 1995, Robins and co-workers purified the enzyme tigloyl-CoA:pseudotropine acyl transferase 330-fold from transformedcultures of D. stramonium. The transferase had a Mr of 65,000,and maximal activity occurred at pH 9 (Rabot et al., 1995). The en-zyme esterifies the 3b-hydroxyl group of pseudotropine (3b-hydroxytropane) with the tiglic acid moiety of tigloyl-CoA. Tropine(3a-hydroxytropane) and norpseudotropine were not acylated.

An early use of 15N NMR spectroscopy in biosynthesis was astudy tracking the metabolic fate of [15N]tropinone in transformedroot cultures of D. stramonium (Ford et al., 1996). Labeling of alka-loids occurred within 2–4 h after administration, particularly to af-ford the tropine esters.

Methyl jasmonate stimulated tropane alkaloid biosynthesis inhairy-root cultures of D. stramonium (Solanaceae) more than ayeast elicitor or oligogalactouronides, and was based on higher lev-els of tropine (Zabetakis et al., 1998).

Contrasting results were observed when a range of analogs of N-methylputrescine and tropinone was fed to transformed root cul-tures of N. rustica and/or a Brugmansia candida � aurea hybrid (Bos-well et al., 1999a). Analogs with larger N-alkyl substituents werenot metabolized. In the N. rustica cultures, the analogs at 1 mMcaused a 4-fold diminution of nicotine (7) in the presence of N-n-propylputrescine. In the B. candida � aurea hybrid culture at1 mM the analogs did not inhibit or substantially interfere withthe accumulation of the normal spectrum of alkaloids.

In follow-up experiments, five enzymes: N-methylputrescineoxidase (MPO), the tropine-forming tropinone reductase (TRI),the pseudotropine-forming tropinone reductase (TRII), the tro-pine:acyl-CoA transferase (TAT), and the pseudotropine:acyl-CoAtransferase (PAT) derived from transformed root cultures of D. stra-monium and a B. candida � aurea hybrid were evaluated for sub-strate specificity (Boswell et al., 1999b). MPO activity was testedwith a series of N-alkylputrescines and N-alkylcadaverines, andTRI and TRII reduction was evaluated against various N-alkylnor-tropinones, N-alkylnorpelletierines and structurally related ke-tones as substrates. TAT and PAT esterification tested a series ofN-substituted tropines, pseudotropines, pelletierinols, and pseudo-pelletierinols as substrates. Some ability of these enzymes to uti-lize unnatural substrates was observed.

Micellar electrokinetic chromatography was used to study thealkaloids scopolamine (11), hyoscyamine (12), and littorine (16)in hairy root clones of Hyoscyamus muticus L. Significant differ-ences were observed in alkaloid content between the clones, andlittorine dominated over scopolamine (11). Aerial parts of H. muti-cus contained no littorine (Mateus et al., 2000).

The mechanism for the conversion of littorine (16), which con-tains a phenyllactic acid ester at C-3 of the tropine nucleus, to atro-pine (12), which has a tropic acid ester moiety, has been a subjectof controversy over many years, since it was reported to occurthrough an intramolecular rearrangement (Ansarin and Woolley,1994). Using (RS)-[2-2H]-, [3,3-2H2]-, and [2,3,3-2H3]phenyllacticacids, Patterson and O’Hagan (2002) showed that the resultingatropine (hyoscyamine) (12) was not formed as a result of a vicinalinterchange taking place during the isomerization. Furthermore, astudy with [10-13C, 30,30-2H2]atropine (12) indicated the stability ofthe alkaloid with no reversible oxidation to the aldehyde occurring.The conclusion was that an S-adenosyl-L-methionine (SAM)/coen-zyme-B12 mediated process was not operating for the isomeriza-tion of littorine (16) to hyoscyamine (12), and that themechanism of the rearrangement remained unknown. Previouswork had examined the stereochemical aspects of the biosynthesis


of tropic acid, present as the esterifying unit of both atropine (12)and scopolamine (11), using specifically labeled phenylalanine pre-cursors (Platt et al., 1984).

Reduction of the 3-keto group in tropinone may occur withdelivery of hydride from the re- or si-face and leads to alkaloidswith a 3a- or a 3b-hydroxy group. These are respectively the atro-pine (12), scopolamine (11) series with a 3a-orientation, or the co-caine (17) and calystegine series which are derivatives ofpseudotropine having a 3b-orientation. Dräger (2006) reviewedthis area of tropane alkaloid enzymology, including aspects of theisolation, cloning, and heterologous expression of the two reduc-tases, TR-I and TR-II, and their protein crystallization and structuredetermination. There is about 50% homology between the two en-zymes, although this may not be meaningful. Also discussed weretheir localization in cultured roots, how they regulate the meta-bolic flux of the tropane alkaloids, and the evolutionary aspectsof the enzymes.

The final step in the biosynthetic pathway for the formation ofscopolamine (11) is epoxidation of 6b-hydroxyhyoscyamine, andan enzyme, 6b-hydroxyhyoscyamine epoxidase, was isolated andcharacterized from cultured roots of H. niger (Hashimoto et al.,1988). The enzyme was a dioxygenase requiring molecular oxygenfor activity.

1.4. Calystegines

The calystegines are a group of hydroxylated nortropane alka-loids with specific glycosidase inhibitory activity. Although it wasinitially thought that their distribution was quite limited, it laterbecame clear that they were also widespread in the Convolvula-ceae (Schimming et al., 1998), which is taxonomically close tothe Solanaceae, a traditional source of tropane alkaloids.

Using 15N-tropinone it was shown in root cultures of Calystegiasepium R.Br. (Convolvulaceae) that pseudotropine (18) and calyste-gine A3 (19) were labeled after 2 days, and calystegines in the Bgroup were labeled subsequently. 2,7-Dihydroxy-nortropane wasalso labeled and maybe a by-product of the tropane pathwaywhich leads to the calystegines (Scholl et al. 2001).

Genes encoding for the formation of the calystegines also occurin plants in the genus Erythroxylum P.Browne (Erythroxylaceae),with alkaloid content up to 0.32% in the leaves. At least one ofthe four main calystegines was found in 38 of 45 herbarium spec-imens of different Erythroxylum species. The family is very distant

taxonomically from the Solanaceae and Convolvulaceae (Brocket al., 2007).

1.5. Pyrrolizidine alkaloids

Homospermidine synthase is the first pathway-specific enzymein pyrrolizidine alkaloid biosynthesis (Böttcher et al., 1993). Rootcultures of Senecio vulgaris L. (Asteraceae) accumulated labeledhomospermidine from putrescine when b-hydroxyethylhydrazine(HEH) was used as a diamine oxidase inhibitor, and was then incor-porated into pyrrolizidine alkaloids when HEH was removed. Theenzyme was partially purified from root cultures of Eupatoriumcannabinum L. (Asteraceae). Putrescine is the exclusive substrate,and the enzyme was inhibited by diaminopropane, spermidine,and cadaverine (mixed inhibition). NADH is also an inhibitor, indi-cating that, even though it is generated in the oxidative deamina-tion of putrescine, it remains enzyme bound. All root cultures ofplants in the Asteraceae which produced pyrrolizidine alkaloidsshowed the presence of homospermidine synthase (Böttcheret al., 1993).

Ober and Kalternegger (2009) reviewed the evolution of the en-zyme homospermidine synthase in plants producing pyrrolizidinealkaloids. It has been shown that the enzyme evolved by duplica-tion of a gene encoding deoxyhypusine synthase, and that this oc-curred several times independently during angiosperm evolution.Only occasionally did these copies function to produce homospe-rmidine synthase, and thus pyrrolizidine alkaloids, indicating thatother regulatory elements were involved in the evolutionaryprocess.

1.6. Retronecine

Bottomley and Geissman (1964) demonstrated that labeledputrescine, and [2- and 5-14C]ornithine, were all precursors ofthe retronecine (20) moiety in pyrrolizidine alkaloids in the rootsof Senecio douglasii DC. (Asteraceae). It was concluded that at leastone of the two molecules of ornithine passes through a symmetri-cal precursor.

An early use of 14C- and 3H-doubly-labeled precursors was anapplication examining the relative rates of incorporation of argi-nine and ornithine into retronecine (20) in Senecio magnificusF.Muell. (Asteraceae). Ornithine was determined to be a signifi-cantly more efficient precursor (Bale and Crout, 1975). Both L-argi-


nine and L-ornithine, but not their D-isomers, served as precursorsof retronecine (20) in Senecio isatideus DC. (Asterceae) (Robins andSweeney 1983).

Senecionine-N-oxide (21) was found to be the primary pyrro-lizidine alkaloid in root cultures of S. vulgaris L. (Asteraceae),after administering a variety of precursors including arginine,ornithine, isoleucine, putrescine, and spermidine. Incorporationof the latter precursors was very high (20–30%) (Hartmann andToppel, 1987). Tertiary pyrrolizidine alkaloids were barelydetected.

Before the development of metabolomics and principal compo-nent analysis, chemical races (chemotypes) within plant specieswere identified by gas chromatography, and subsequently by gaschromatography coupled with mass spectrometry (GC–MS). Exam-ination by GC–MS of more than 100 populations of Senecio jacobaeaL., and about 25 populations of S. erucifolius L., from different geo-graphical locations for the content of sixteen pyrrolizidine alka-loids established the presence of different chemotypes (Witteet al., 1992). A jacobine chemotype and an erucifoline chemotypewere observed for S. jacobaea, and an erucifoline chemotype andan eruciflorine chemotype for S. erucifolius. Within the respectivespecies there were no morphological differences, however, ob-served between the chemotypes. Eruciflorine (21-hydroxyinteg-errimine) was identified as a new alkaloid.

For the first time, in vivo 15N NMR spectroscopic studies wereused to study alkaloid metabolism in D. stramonium and N. taba-cum transformed root cultures (Ford et al., 1994). The amount ofdetectable 15N label in the secondary metabolites was about1.4 lmol. In D. stramonium, the 15N resonance of hyoscyamine(12) was observed at �305.5 ppm, along with three unidentifiedalkaloids, and possibly some conjugated putrescine derivatives.In N. tabacum, nicotine (7) showed 15N resonances at �75.7 and�309 ppm for the pyridine and pyrrolidine nitrogen atoms, respec-tively. Aspects of the subcellular distribution of nicotine (7) werededuced.

If alkaloids are viewed as protective agents, then the most vul-nerable parts should be the most protected. The youngest leaves ofCynoglossum officinale L. (Boraginaceae) contained up to 190 timeshigher levels of pyrrolizidine alkaloids than the older leaves (vanDam et al., 1994). Similar defensive effects were observed for thequinolizidine alkaloids of Spanish broom, Spartium junceum L.(Fabaceae), where the highest alkaloid contents were found inyoung plants, in reproductive organs, such as the seeds, and in pri-mary twigs (Barboni et al., 1994).

Theoretical considerations suggest that incorporation of (S)-[1-2H]-putrescine through the symmetrical intermediate homo-spermidine into the necine unit of the pyrrolizidine alkaloidsshould proceed with 50% deuterium retention. However, experi-mentally a level of only 34% to 34.5% retention was found intwo different laboratories. Graser and co-workers (1998) con-firmed these results and eliminated deuterium isotope effectsduring homospermidine formation as an explanation. Using S.

vulgaris root cultures, 2H-retention for putrescine, spermidine,and homospermidine was examined after administration, dou-bly-labeled [2H-14C]putrescine. Stereospecific loss from (S)-[1-2H]-labeled putrescine occurred during the conversion of sper-midine into putrescine during its reversible interconversion,accounting for the difference between predicted and determined2H-retention. The mechanism of the reaction remained to beelucidated.

In Crotalaria scassellatii Chiov. (Fabaceae) seeds, pyrrolizidinealkaloids are stored as tertiary amines. At seed germination, con-version to the alkaloid N-oxides transportation, metabolism, andstorage occurs. A flavin-dependent, mixed function N-oxygenase(SNO) was isolated and partially purified. It showed low Km valuesfor senecionine and monocrotaline (Chang and Hartmann, 1998).The substrate specificity was very similar to an insect SNO charac-terized from the hemolymph of arctiid larvae (Lindigkeit et al.,1997).

A further study of 24 species of the Jacobaea section of thegenus Senecio for the presence of 26 pyrrolizidine alkaloids(PAs) indicated that, with only one exception, all were derivativesof senecionine (22). Based on this, a biosynthetic grid was devel-oped involving two pathways, and an evolutionary history wasdeveloped. It was concluded that the differences in PA profiles be-tween the species, and in other sections of Senecio, can be attrib-uted to transient switching of the genes encoding the PApathway-specific enzymes, not their presence or absence (Pelseret al., 2005).

1.7. Lysine-derived alkaloids

The presence of lysine decarboxylase was demonstrated to bepositively correlated with leaf alkaloid content in Lupinus polyphyl-lus Lindl. (Fabaceae), and the enzyme was demonstrated to bepresent in 46 alkaloidal and non-alkaloidal species in 17 plant fam-ilies (Schoofs et al., 1983).

Quinolizidine alkaloid distribution from seed germination toseed formation was studied in three Lupinus species, and discussedin relationship to predation (Williams and Harrison, 1983). Alka-loid levels were highest at the time of seed formation, and declinedrapidly after germination.

Quinolizidine alkaloids are mainly distributed in the subfamilyPapilionoideae in the family Fabaceae, and a well-establishedsource is the genus Lupinus. The first plant-derived lysine decar-boxylase activity was reported from Lathyrus sativus L. (Fabaceae)(Ramakrishna and Adiga, 1976). There is about a four-fold differ-ence in the alkaloid levels between the sweet and the bitter lupinsLupinus luteus L. and L. albus L. (Fabaceae). It was deduced thatcyclization steps in the quinolizidine pathway were blocked inthe sweet lupins, although acyltransferase levels were the samein both lupin varieties (Saito et al., 1993).

In a subsequent study, the sweet and bitter varieties of threeLupinus species (L. angustifolius L., L. albus, and L. luteus) quino-lizidine alkaloid levels varied from 0.58 lg/ml (sweet) to29.6 lg/ml (bitter) in L. albus (Aniszewski et al., 2001). Bitter lu-pin seeds also contained high levels of biogenic polyamines, andlow basic amino acid levels. The reverse was noted for sweet lu-pin seeds.

The subcellular localization of two acyltransferases was stud-ied in L. albus and L. hirsutus L. The enzymes were not locatedin the chloroplasts where de novo alkaloid synthesis occurs. Onewas discovered in the mitochondrial fraction and localized inthe matrix, and the other was in the cytosol. A proposal for thedetailed transport of quinolizidine alkaloids within the cell basedon the location of the two enzymes was made (Suzuki et al.,1996).


1.8. Phenylalanine–tyrosine-derived alkaloids

1.8.1. Benzylisoquinolines and derivativesOne of the most important alkaloid biosynthesis papers pub-

lished in Phytochemistry in the 50-year period was that by Zenkand co-workers (Stadler et al., 1989) on the role of (S)-norcoclaurine(23) in benzylisoquinoline alkaloid biosynthesis. Based on long-held

biogenetic ideas and precursor administration experiments by oth-ers, it was considered that the tetrahydroxylated norlaudanosoline(24) was the precursor of the complex benzylisoquinoline alkaloids.However, Zenk and co-workers demonstrated that (S)-[1-13C]norco-claurine (23), a trihydroxylated intermediate, was a precursor of pa-vine, protoberberine, aporphine, and benzophenanthridinealkaloids in selected cell cultures and whole plants. This dramaticresult revised the early steps in benzylisoquinoline alkaloid biosyn-thesis showing that tyrosine is metabolized to dopamine for theupper unit, and then condenses with 4-hydroxyphenylacetaldehydeto form 23. This result also explains why dopamine, or related 3,4-dihydroxy derivatives, were not incorporated into the lower unitsof benzylisoquinoline alkaloids.

The step after O- and N-methylation in the benzylisoquinolinebiosynthesis pathway is hydroxylation of ring C. Loeffler and Zenk(1990) reported on the isolation of an enzyme purified 488-foldfrom cultures of Berberis stolonifera Koehne & E.L.Wolf (Berberida-ceae) which catalyzed the meta-hydroxylation of coclaurine (25),as well as N-methylcoclaurine, tyrosine, and tyramine.

Several isoquinoline-containing species belonging to the fami-lies Berberidaceae, Fumariaceae, Papaveraceae, and Ranunculaceaeafforded an enzyme complex SAM:30-hydroxy-N-methyl-(S)-cocla-urine-40-O-methyltransferase (40-OMT) (Frenzel and Zenk, 1990b).The enzyme was purified 400-fold from Berberis koehneana Schnei-der (Berberidaceae) cell cultures, and consisted of only the 40-OMTand SAM:norcoclaurine-6-O-methyltransferase; further separationwas not successful. The 40-OMT was substrate specific, and, in thepresence of SAM, catalyzed methyl transfer to the C-40 hydroxylgroup of benzylisoquinoline alkaloids with the 1S-configurationand having ortho hydroxy groups at C-30 and C-40; the natural sub-strate is 30-hydroxy-N-methyl-(S)-coclaurine (26). The pH opti-mum was 8.3, it had Mr 4.0 kD, and was potently inhibited bythe product alkaloid. The half life of the enzyme was three daysat room temperature, and it was stable for >1 year at �20 �C.

Morphine (27) biosynthesis requires a reduction step after theoxidation of (S)-reticuline (29) to afford (R)-reticuline (28), andthe NADPH-dependent enzyme, 1,2-dehydroreticuline reductase,was found in seedlings of Papaver somniferum (De-Eknamkul andZenk, 1992); it was not found in non-morphine containing plants.

The isolated cytosolic enzyme had a Mr of 30 kD, a pH optimum of8.5 and a temperature optimum of 30 �C. The Km value for 1,2-dehydroreticuline (30) was 10 lM. The enzyme, which transfersthe pro-S-hydride of NADPH to C-1 of 1,2-dehydroreticuline (30),shows high substrate specificity, since neither 1,2-dehydronorret-iculine nor 1,2-dehydrococlaurine were substrates; both (S)- (29)and (R)-reticuline (28) inhibit the enzyme.

Peumus boldus Molina (Monimiaceae) and B. stolonifera Koehneet E.L.Wolf (Berberidaceae) are sources of aporphines and protob-erberines, respectively. When triply-labeled [1-13C,6-O13CH3,N-13CH3]-(S)-reticuline (29) was administered to their cellcultures unanticipated transmethylations of the methyl groups in


both aporphines and protoberberines were observed by 13C NMRspectroscopy. The phenomenon was explained by demethylation,incorporation into the C-1 pool, and remethylation (Schneiderand Zenk, 1993a).

When 13C-labeled (S)-reticuline (29) was utilized in a non-alka-loid producing cell line of Thalictrum tuberosum L. (Ranunculaceae),the protoberberine alkaloid scoulerine (31) was labeled initially,followed by other protoberberine derivatives (Schneider and Zenk,1993b). This suggested that early stage events in the pathway weredownregulated in the cell line.

From cultures of Thalictrum minus L. (Ranunculaceae) S-adeno-syl-L-methionine:norcoclaurine 6-O-methyltransferase was puri-fied and characterized (Hara et al., 1995). In 1995, Zenk and co-workers (Loeffler et al., 1995) showed that norcoclaurine (23) inthe presence of S-adenosyl-L-methionine (SAM) was transformedinto N-methylnorcoclaurine, coclaurine (25), and N-methylcoclau-rine. Eight plant cell cultures [Dicentra spectabilis Lem. (Fumaria-ceae), Fumaria capreolata Leight. (Fumariaceae), Chondodendrontomentosum Ruiz & Pav. (Menispermaceae), Tinospora cordifoliaMiers (Menispermaceae), Argemone platyceras Link & Otto (Pap-averaceae), P. somniferum, B. stolonifera, and Berberis julianaeC.K.Schneid. (Berberidaceae)] were studied. In T. cordifolia, (S)-nor-coclaurine (23), but not its (R)-enantiomer, was exclusively O- andN-methylated. The enzyme responsible for the N-methylation waspurified and shown to catalyze stereoselectively only the methyltransfer from SAM to (S)-configured norcoclaurine (23) and cocla-urine (25). Thirteen other benzylisoquinoline alkaloids wereunsuccessfully used as substrates. The pH optimum was 8.6, andas a substrate, (S)-coclaurine (25) showed a Km of 36 lM; themolecular weight of the enzyme was estimatd as 85 kD. It was de-duced that the stereoselective enzyme was present in only certainmembers of the Menispermaceae and in D. spectabilis Lem.(Fumariaceae).

The occurrence of (S)-norcoclaurine synthase (NCS) in ninetyspecies of plants suggested the monophyletic origin of benzyliso-quinoline alkaloid biosynthesis prior to the emergence of the eudi-cots. A latent molecular fingerprint for benzylisoquinolinebiosynthesis in plants not known to produce such alkaloids wasalso detected based on analysis for NCS, berberine bridge enzyme,and several O-methyltransferases (Liscombe et al., 2005).

As mentioned, for many years, norlaudanosoline (24) was re-garded as the immediate precursor for many benzylisoquinolineand related alkaloids. Papaverine (32) was an early speculatedproduct thought to be derived through complete O-methylationof the tetrahydroxylated intermediate [norlaudanosoline (24)]and aromatization. Following more detailed precursor adminis-tration studies, Zenk and co-workers completely revised thispathway (Han et al., 2010). It was shown using 8-day-old seed-lings of Papaver followed by direct examination of the stable iso-tope labeled precursors in the total plant extract, that thetrihydroxylated norcoclaurine (23) is the primordial benzyliso-quinoline alkaloid, and is further converted into (S)-reticuline(29). Methylation of (S)-reticuline (29) generates (S)-laudanine,and methylation at the 30-position leads to laudanosine (33),known alkaloids from the opium poppy. N-Demethylation of lau-danosine (33) then affords 1,2,3,4-tetrahydropapaverine. Dehy-drogenation is step-wise with 1,2-dihydropapaverine as anintermediate, followed by dehydrogenation of the 1,2-bond topapaverine (32); the mechanism of the process is not known.The previously claimed nor-reticuline (34) was not involved inpapaverine (32) biosynthesis.

Studies on the relationships of the proaporphine alkaloids,such as crotonosine (35), and the dihydroproaporphine alkaloids,such as linearisine (36), became somewhat more confused whenit was shown that linearisine (36) can be oxidized, undergoinversion at C-6a, and N-demethylation to afford crotonopsine

(35) in Croton linearis Jacq. (Euphorbiaceae) (Stuart and Graham,1973b).

1.9. Bisbenzylisoquinoline alkaloids

Very few studies have been reported on the biosynthesis of thebisbenzylisoquinoline alkaloids. In 1980, Bhakuni and co-workers(1980) reported on the incorporation of doubly-labeled (±)-N-methylcoclaurine (37) into tetrandrine (38), and showed that theC-1H was retained.

When 14C-labeled tyrosine, tyramine, and several chiral 1-ben-zyl-1,2,3,4-tetrahydroisoquinolines were administered to bisben-zylisoquinoline alkaloid-producing cell cultures of B. stolonifera(Stadler et al., 1988), berbamunine (39) showed significantly high-er incorporation than 2-norberbamunine, berbamine, aromoline,or isotetrandrine. This suggesed that 39 is possibly the initial dimerformed. Exclusive incorporation of tyramine into the isoquinolineportion of this dimer was observed. Administering various benzyl-isoquinoline precursors showed that (R)-N-methylcoclaurine (37a)gave the highest relative incorporations (29% into berbamunine)and (S)-N-methylcoclaurine (37b) the lowest (2.4%). Both (S)-coclaurine (25) and (S)-N-methylcoclaurine (37b) were well incor-porated into the protoberberine fraction (21 and 54%, respectively).Consequently, the pathway which promotes dimerization and theC-30-hydroxylation pathway leading to reticuline are in competi-tion for these substrates. Using [1-13C]-(R)- and (S)-coclaurine(25a and 25b) showed that the (R)-isomer (25a) is incorporatedsimilarly into both halves of the (R, R)-dimer guattegaumerine.Racemization of (S)- to (R)-coclaurine did not occur.

1.10. Berberines and derivatives

An N-methylation step, followed by hydroxylation a to N, is in-volved in the biosynthesis of the benzazepine alkaloid cis-alpinig-enine (40). When [8,13,14-3H3]-(±)-tetrahydropalmatine (41) and[8,13,14-3H3,8-14C]-(±)-tetrahydropalmatine methiodide (42) wereadministered to Papaver bracteatum the same loss of 3H was ob-served, including complete 3H loss from C-14 (Roensch, 1977).The next alkaloid in the biogenetic sequence, [8-14C]muramine(43), was efficiently incorporated into alpinigenine (40). Usingunnatural substrates, such as [20-14C]hydroxymethylaudanosineand [8-14C]13-hydroxymuramine (44), the specificity of alkaloidmetabolism was examined. Tracer dilution was used to confirmthe occurrence of the three established intermediates.

The most important intermediate enzyme in the formation ofmany advanced benzylisoquinoline alkaloids is the berberinebridge enzyme (BBE) which catalyzes the oxidation of a N-methylgroup, the coupling to form the C-ring of protoberberine, and sub-sequently the benzo[c]phenanthridine alkaloids. Zenk and co-workers (Steffens et al., 1985) found the enzyme in 66 samplesof differentiated plants and cell suspension cultures. The enzymewas purified 450-fold from Berberis beaniana C.K.Schneid. (Berbe-ridaceae) cell cultures and shown to be homogeneous by gel elec-trophoresis (mol. wt. 52 kD). The enzyme required the presence ofoxygen and catalyzed the transformation of the (S)-enantiomers ofreticuline (29), protosinomenine, and laudanosoline to the corre-sponding (S)-tetrahydroprotoberberines, releasing stoichiometricamounts of H2O2 in the process.

The STOX enzyme, (S)-tetrahydroprotoberberine oxidase, wasenriched 7.4-fold from a high-yielding cell suspension of Berberiswilsoniae Hemsl. var. subcaulialata and immobilized (Amann andZenk, 1987). Compared with immobilized Berberis cells and withthe soluble enzyme, the pH and temperature optima were slightlyshifted by immobilization, but the Km values for (S)-norreticuline(34a) were unaffected. Stability was improved 50-fold comparedwith the free enzyme. A cyclic C-1 inversion process was described


using the stereospecific enzymatic oxidation of (S)-norreticuline(34a) to 1,2-dehydronorreticuline followed by sodium borohydridereduction to the (R)-enantiomer (34b).

The role of protopine (40) in benzo[c]phenanthridine biosynthe-sis was defined by Tanahashi and Zenk (1990). [6-3H]Protopine (45)was synthesized, and in Eschscholzia californica suspension culturesinduced by a yeast preparation, produced 6-hydroxyprotopinewhich spontaneously rearranged to [11-3H]-7,8-dihydrosanguinar-ine. It was deduced that the enzyme was a P450-linked monooxy-genase which could be induced by elicitor and was present indifferent plant species producing benzo[c]phenanthridine alkaloids.

An enzyme was found in different species of isoquinoline alka-loid-producing plant cell cultures which specifically N-methylatescertain (S)-tetrahydroprotoberberine alkaloids, such as (S)-cana-dine (46) and (S)-stylopine (47), at the expense of S-adenosyl-L-methionine (SAM). It was partially purified (90-fold) from E. cali-fornica Cham. (Papaveraceae) cell suspension cultures and charac-terized. The enzyme had a pH optimum of 8.9, a temperatureoptimum of 40 �C, and a molecular weight of 78 kDa. The Km for(S)-canadine (46 was 6.4 lM, and for (S)-stylopine (47) was3.1 lM, and the enzyme was inhibited by S-adenosyl-L-homocyste-ine (SAH) with a Ki of 24 lM (Rueffer et al., 1990).

After screening 37 cell suspension cultures and finding N-meth-yltransferase activity in 34 of them, B. koetineana, which producedthe highest enzyme levels, was selected for further study. Threedistinctly different isoforms of S-adenosyl-L-methionine-(R,S)-tet-rahydro-benzylisoquinoline-N-methyltransferases were isolatedand designated as NMT-I, -II, and -III. The three enzymes haddifferent Mr of 60–78 and 103 kD, pH optima (6.8/7.4), kinetic

properties, and substrate specificities. NMT-I showed maximalactivity with (R)-tetrahydropapaverine as substrate, whereasNMT-II and -III were most active against (R)-coclaurine (25a)(Frenzel and Zenk, 1990a). After immobilization, the enzymes re-tained their properties, and may serve for the preparation of isoto-pically labeled N-methylated benzylisoquinoline alkaloids.

The enzymatic synthesis of the methylenedioxy moiety in manybenzylisoquinoline alkaloid derivatives had long been a subject ofdiscussion and experimentation; Bauer and Zenk (1991) resolvedthis issue in an elegant manner. A microsomal preparation fromE. californica cell suspension cultures in the presence of NADPHand O2 catalyzed the formation of two methylenedioxy bridges intwo consecutive steps: (S)-[10-OC3H3]scoulerine (31) afforded(S)-cheilanthifoline and (S)-[3-OC3H3]cheilanthifoline produced(S)-stylopine (47). Nandinine was not an intermediate. The releaseof HO3H into the aqueous incubation mixture was used as an assayfor both enzyme activities. The enzymes were characterized ascytochrome P450 monooxygenases on the basis of substrate spec-ificity, cofactor requirements, and inhibition by cytochrome P450specific inhibitors. The enzymes were induced (12- and 5-fold)20 h after challenging the cell suspension culture with elicitor.The methylenedioxy moiety is viewed as derived from the oxida-tion of an o-methoxyphenol.

From several different Ranunculaceae and Berberidaceae cellcultures, an enzyme system catalyzing methylenedioxy bridgeformation on ring A of (S)-canadine [(S)-tetrahydroberberine](46) from (S)-tetrahydrocolumbamine was found (Rueffer andZenk, 1994). The cytochrome P450 enzyme complex waspartially characterized from a protoberberine-alkaloid producing


T. tuberosum L. (Ranunculaceae) cell line, and consisted of amicrosomal associated oxidase and a cytochrome P450 reduc-tase with a pH optimum at pH 8.5 and a temperature optimumof 40 �C. The apparent Km value was 11.5 lM fortetrahydrocolumbamine.

1.11. Benzo[c]phenanthridines

In suspension cultures of E. californica, the benzo[c]phenanthri-dine alkaloids sanguinarine (48) and chelerythrine (49) reachedmaximum levels a few hours after elicitation by a yeast extract.Subsequently, their levels declined, and the level of macarpine(5-methoxychelirubine) (50), the most highly oxidizedbenzo[c]phenanthridine alkaloid, increased. Induced tyrosinedecarboxylase activity correlated with alkaloid formation (Collingeand Brodelius, 1989).

Temporal studies correlating the metabolism of tyramine withthe production of sanguinarine (48) and hydroxycinnamic acidamides in P. somniferum cultures showed that elicitors tended toinduce tyramine hydroxycinnamoyl transferase, which led to highlevels of the amides in cell wall fractions, and only a small increasein alkaloid production through induction of berberine bridge en-zyme (Facchini, 1998).

Dihydrosanguinarine-10-hydroxylase and 10-hydroxydihydro-sanguinarine-10-O-methyltransferase, two new enzymes involvedin benzo[c]phenanthridine alkaloid biosynthesis, were found incell-free extracts derived from E. californica cell suspension cultures(De-Eknamkul et al. 1992). The hydroxylase is a microsome-associ-ated, cytochrome P450-dependent monooxygenase which acts spe-cifically at C-10 of 7,8-dihydrosanguinarine (51). The

methyltransferase, which was enriched 60-fold and partially char-acterized, exclusively methylates the C-10 hydroxyl moiety to formdihydrochelirubine (10-methoxy-7,8-dihydrosanguinarine) (52).High substrate specificity was observed for both enzymes.

Cell-free extracts of yeast-elicited Thalictrum bulgaricum Velen.(Ranunculaceae) cell cultures afforded two novel enzymes, dihydr-ochelirubine-12-hydroxylase and SAM:12-hydroxydihydrocheliru-bine-12-O-methyltransferase which are involved in the last twosteps in macarpine biosynthesis (Kammerer et al., 1994). Thehydroxylase was a microsomal-associated, cytochrome P450-dependent monooxygenase acting exclusively at C-12 of dihydr-ochelirubine (52). The methyltransferase acted highly specificallyon the hydroxyl moiety at C-12 to form 7,8-dihydromacarpine(53). As a result, the complete sequence of macarpine (50) biosyn-thesis was clarified at the enzymic level, the longest alkaloid path-way at that time.

An elicitor derived from Penicillium expansum, or the calciumionphore A23187, induced benzo[c]phenanthridine alkaloid pro-duction in Sanguinaria canadensis L. (Berberidaceae) suspensioncultures. Adding either a calcium chelatant or a calcium channelinhibitor, such as verapamil, markedly decreased alkaloid produc-tion, suggesting that calcium ions were important for stimulatingalkaloid biosynthesis (Mahady and Beecher, 1994).

Further studies showed that compounds which modify proteinkinase activity, such as phorbol-12-myristate-13-acetate and 1-oleoyl-2-acetyl-glycerol, significantly (65-fold and 25-fold, respec-tively) increased benzo[c]phenanthridine alkaloid production,whereas a protein kinase C inhibitor, such as staurosporine, mark-edly decreased alkaloid production in S. canadensis cell cultures(Mahady et al., 1998). G-Protein activators, such as mastoparan,


mas-7, or melittin also induced alkaloid biosynthesis indicatinginvolvement in signal transduction.

1.12. Morphinan and related alkaloids

An early evaluation of the enzymatic capacity of the latex of P.somniferum L. (Papaveraceae) was a demonstration that a centri-fuged preparation was capable of converting dihydroxyphenylala-nine into morphine (27) in vitro (Fairbairn and Djote, 1970);further metabolism of morphine (27) was noted in the stem latex,but not the capsule latex.

The morphinandienone alkaloid sinoacutine (54) is in the enan-tiomeric series compared with morphine (27). An early detailedstudy of alkaloids in this series using 14C- and 3H-labeled alkaloidprecursors was reported by Stuart and Graham (1973a).

[3-14C]-1,2-Dehydroreticulinium (30) and [3-14C]-(±)-reticuline(28/29) were incorporated into thebaine in P. bracteatum Lindl. (Pap-averaceae). [16-3H]Codeinone (55) was reduced to codeine (56), butdemethylation reactions were not observed (Hodges et al., 1977).

Roberts and Antoun (1978) detected L-dopa decarboxylase inpoppy latex using 14C-labeled dopa derivatives as substrates. Max-imum activity was observed at pH 7.2, and L-tyrosine, L-phenylala-nine, and L-histidine were also substrates.

In cell cultures of P. somniferum (±)-reticuline (28/29) was stereo-specifically converted to (�)-(S)-scoulerine (31) and (�)-(S)-chei-lanthifoline. (�)-Codeinone (55) was converted in high yield to(�)-codeine (56) in both cell culture and enzyme preparations; the-

baine (57), codeine (56), and morphine (27) were not metabolized(Furuya et al., 1978). Codeine (56) was converted to morphine (27)in isolated capsules of P. somniferum; co-factors such as NAD, ATP,S-acetyl CoA, and pyridoxal phosphate were not required (Hsuet al., 1985).

The conversion of (R)-reticuline (28) to salutaridine (58) wascatalyzed by microsomal preparations derived from differentiatedP. somniferum plants or a thebaine-producing cell suspension cul-ture (Gerardy and Zenk, 1992). NADPH is a required co-factor, asis molecular oxygen, and based on inhibition experiments withnaphthoquinones, ancymidole, and prochlorax, the enzyme is acytochrome P450-dependant oxidase which refuted earlier claimsabout the formation of this key intermediate from (R)-reticuline(28). The enzyme is present in the roots, shoots and capsules ofthe poppy plant.

Gerardy and Zenk (1993) also isolated the cytosolic enzyme sal-utaridine 7-reductase from P. somniferum cell cultures and differ-entiated plants. The enzyme, of approximate molecular weight52,000 daltons, and a pH optimum of 6.0–6.5, stereoselectively re-duces salutaridine (58) to (7S)-salutaridinol (59) using NADPH ascosubstrate. Screening of isoquinoline-containing plants and cellcultures showed that the enzyme occurs only in P. somniferumand P. bracteatum.

[N-14CH3]Thebaine (57), when applied to a low-thebaine yield-ing cell suspension culture of P. somniferum, afforded codeine (56)and two new metabolites, tetrahydrothebaine and thebainone,showing that poppy cell cultures contain the enzymes necessary


for thebaine enolether cleavage at C-6 (Wilhelm and Zenk, 1997).When the ability of 60 cell suspension cultures of different plantspecies and 230 field grown plant species was evaluated for theircapability to metabolize [N-14CH3]thebaine (57), only the singlecell culture of Mahonia nervosa Nutt. (Berberidaceae) cleaved theC-3 phenol methylether to yield oripavine. Field-grown M. nervosacould not perform this reaction (Wilhelm and Zenk, 1997).

Thirty years after the initial studies, the biological scene wasvery different when Huang and Kutchan (2000) reported on themorphinan and benzo[c]phenanthridine alkaloid gene transcriptaccumulation in P. somniferum. Using RNA gel blot analysis, thetranscripts of three genes of alkaloid biosynthesis in P. somniferumwere monitored in developing seedlings, mature plants, and plantcell suspension culture. The genes encoded the early enzyme (S)-N-methylcoclaurine 30-hydroxylase (cypb80b1) which that is com-mon to both morphine and sanguinarine biosynthesis, the berber-ine bridge enzyme (BBE) which lies on the pathway tosanguinarine (48), and codeinone reductase (COR), the penultimateenzyme in the biosynthesis of morphine (27) biosynthesis. Theba-ine (57) was present throughout the 20-day germination period,but sanguinarine (48) was only detected after day 5, which paral-leled the accumulation of the cyp80b1, bbe1, and cor1 gene tran-scripts. In the mature plant, the cyp80b1, bbe1 and cor1 genetranscripts were detected in the root, the stem, the leaf lamina,and the leaf mid rib. However, only cyp80b1 and cor1 were foundin the flower bud and the capsule, paralleling the absence of san-guinarine (48). The cyp80b1 and bbe1 gene transcripts were dem-onstrated in cell suspension cultures induced with methyljasmonate, consistent with the presence of sanguinarine.

Facchini and Park (2003) demonstrated the significant homol-ogy between the clones of three methyltransferases involved in(S)-reticuline (29) biosynthesis, (S)-norcoclaurine-6-O-methyl-transferase, (S)-30-hydroxy-N-methylcoclaurine-40-O-methyltrans-ferase, and (S)-coclaurine-N-methyltransferase, in P. somniferumand Coptis japonica Makino (Ranunculaceae). These cDNAs wereused with clones for tyrosine/dopa decarboxylase, (S)-N-methylco-claurine-30-hydoxylase, berberine bridge enzyme, (7S)-salutaridi-nol 7-O-acetyltransferase, and codeinone reductase, to examinethe accumulation of gene transcripts for alkaloid production in P.somniferum. With the exception of codeinone reductase, all of theenzymes were induced by an elicitor or by wounding suggestinga coordination of both developmental and inducible regulation ofthe genes for alkaloid biosynthesis (Facchini and Park, 2003).

In the morphine biosynthesis pathway, inversion of configura-tion at C-1 is required in reticuline from (S)- (29) to (R)- (28).Zenk’s group showed that seedlings of P. somniferum containedan unstable synthase in the vesicles which oxidizes (S)-reticuline(29) to 1,2-dehydroreticuline; (S)-norreticuline was also a sub-strate. It was partially purified and had a pH optimum of 8.75, withan apparent Km value for reticuline of 117 mM. In combinationwith the previously discovered 1,2-dehydroreticuline reductase,the 1,2-dehydroreticuline synthase provided for the requiredinversion of configuration, and completed the pathway of elevenenzymes, beginning with two molecules of L-tyrosine via (S)-nor-coclaurine (23) to (R)-reticuline (28) (Hirata et al., 2004).

Further studies by the same group, using [ring-13C6]-L-tyrosine,[ring-13C6]-tyramine, and [1,2-13C2], [6-O-Me 13C]-(±)-coclaurine(25), elucidated all the major fragment ions of labeled codeine(56) and morphine (27) through a combination of LC-electrosprayMS/MS and high resolution FTICR-MS (Poeaknapo et al., 2004).

An important and new approach to studying the range of alka-loids labeled by a given precursor, and an easier method to assessthe flux of alkaloids, was reported by Zenk and co-workers(Schmidt et al., 2007). Using [ring-13C6]-tyramine as a precursorof the alkaloids of P. somniferum, the labeling into twenty charac-terized alkaloids of the morphinan, benzylisoquinoline, protoberb-

erine, benzo[c]phenanthridine, phthalide-isoquinoline, andprotopine types was described. The alkaloid patterns were deter-mined by direct infusion high-resolution ESI-FTICR mass spectrom-etry and HPLC/electrospray tandem mass spectrometry.

On the morphine biosynthetic pathway, salutaridine (58) is ste-reospecifically reduced by salutaridine reductase (SalR) to 7S-salutaridinol (59), which is then acetylated by salutaridinol 7-O-acetyltransferase (SalAT). P. somniferum was transformed with anRNAi construct designed to reduce transcript levels of the geneencoding the SalAT, which led to the accumulation of salutaridine(58) and salutaridinol (53) (Kempe et al., 2009). Using quantitativePCR, the SalAT transcript was shown to be diminished, while salRtranscript levels were unaffected. Concentrations of thebaine (57)and codeine (56) in the latex were reduced, while morphine (27)levels remained constant.

Ziegler and co-workers (2009) reviewed aspects of the evolu-tion of morphine (27) biosynthesis, since this is a rare biosyntheticpathway compared with all the plants producing benzylisoquino-line alkaloids (BIA). Comparative transcriptome analysis of opiumpoppy and Papaver species which do not accumulate morphinanalkaloids showed that known genes encoding BIA biosynthetic en-zymes are expressed at higher levels in P. somniferum. Salutaridinereductase (SalR) was isolated and heterologously overexpressed.Homologous modeling and substrate docking established the sub-strate binding site for SalR which was compared to several mem-bers of the short chain dehydrogenase/reductase (SDR) familyfrom different plant species.

1.13. Miscellaneous alkaloids

1.13.1. SalsolineRadiolabeled 3-hydroxy-4-methoxyphenylethylamine and 6-

hydroxy-7-methoxy-1-methyl-1,2,3,4-tetrahydroisoquinoline-1-carboxylic acid were precursors of salsoline (60) in Echinocereusmerkeri Hildm. ex K.Schum. (Cactaceae) (McFarlane and Slaytor,1972).

1.13.2. GalanthamineZenk and co-workers showed that administering 13C-labeled 40-

O-methylnorbelladine (61) resulted in high levels of incorporationinto galanthamine (62) (27%) and N-demethylgalanthamine (63)(31%) in cultivated Leucojum aestivum L. (Amaryllidaceae) (Eich-horn et al. 1998). In addition, 40-O-methylbelladine (64) was a poorprecursor, and it was shown that the final step in galanthaminebiosynthesis was N-methylation. As a result, a new scheme forgalanthamine biosynthesis was proposed in which phenol oxida-tive coupling of 40-O-methylnorbelladine (61) is the first step.

1.13.3. OnychineA simple, novel azafluorene alkaloid onychine (65) was charac-

terized from the trunk wood of Onychopetalum amazonicum R.E.Fr.(Annonaceae) (De Almeida et al., 1976), after the original isomericstructure (1-aza-4-methylfluoren-9-one) was demonstrated bysynthesis to be incorrect. The alkaloid was subsequently isolatedby Waterman and Muhammad (1985) from Cleistopholis patensEngl. & Diels (Annonaceae), together with the 1-aza-4-methylan-thraquinone alkaloid, cleistopholine (66), and the oxoaporphinealkaloid liriodenine. Cavé and co-workers (Tadic et al., 1987) char-acterized another member in the series kinabaline (5,8-dimethoxy-6-hydroxyonychine) (67), from the trunk bark of Meiogyne virgataMiq. (Annonaceae), and extended an earlier proposal regarding thebiogenesis of the azaanthracene and the azafluorene alkaloids fromoxoaporphinoid precursors.


1.13.4. SecurinineThe alkaloid securinine (68) represented a new alkaloid skele-

ton when it was first isolated in 1963. Biogenetically, it wasthought to arise in part from lysine, but the origin of the remain-ing carbons and oxygens was unclear for many years. In 1977,the breakthrough came when Sankawa and co-workers (1977)demonstrated that lysine, cadaverine, and tyrosine were incorpo-rated into 68 in Securinega suffruticosa (Pall.) Rehder (Euphorbia-ceae). DL-[2-14C]Tyrosine specifically labeled the lactone carbonylgroup, and using L-[U-14C]tyrosine and L-[30,50-3H2]tyrosine it wasshown that the aromatic ring and C-2 and C-3 of tyrosine wereincorporated intact into 68. Similar to the alkaloid profile of theroots, allosecurinine (69) (major) and securinine (68) were pro-duced in callus cultures derived from budding stem explants ofthe seeds of S. suffruticosa, and the factors affecting alkaloid pro-duction (pH, hormones, light/dark) were examined (Ide et al.,1986).

1.13.5. ColchicineSignificantly improved parameters to enhance incorporation

rates for the study of colchicine (70) biosynthesis in immature seedsof Colchicum autumnale L. (Colchicaceae) were developed by Zenkand co-workers (Nasreen et al., 1997). Using 3H- and 13C-labeledphenethylisoquinolines, it was shown that N-methylation occursprior to substitution of ring C, and that a C-13 hydroxy group inautumnaline (71) is necessary for phenolic coupling reaction.Autumnaline (71) afforded a broad range of metabolic products inthe seeds.

1.13.6. Erythrina alkaloidsZenk’s group also developed an improved delivery system of

radio- and stable isotope precursors for Erythrina alkaloids in Ery-

thrina crista-galli L. (Fabaceae), following the demonstration thatthe fruit wall tissue was the main site of alkaloid biosynthesis (Ma-ier et al., 1999). This ground-work led to the discovery that the pre-viously assumed precursor, (S)-norprotosinomenine (72), was notincorporated into the Erythrina alkaloids. However, (S)-coclaurine(25b) and (S)-norreticuline (34a) were metabolized to erythraline(73) and erythrinine (74), respectively. [1-13C]-(S)-Norreticuline(34a) exclusively labeled erythraline (73) at position C-10, indicat-ing that the participation of a symmetrical intermediate in Erythri-na alkaloid biosynthesis could be excluded.

1.13.7. Ipecac alkaloidsThe ipecac alkaloids are comprised of a tetrahydroisoquinoline

moiety linked to a monoterpenoid unit. De-Eknamkul et al.(1997) found two enzyme activities in cell-free extracts of theleaves of Alangium lamarckii Thwaites (Alangiaceae). The first stepin the pathway, condensation of dopamine and secologanin,

occurred rapidly at pH 7.5 to produce (1R)-deacetylipecoside (75)and (1S)-deacetylisoipecoside (76) which spontaneously convertedto demethylalangiside (77) and demethylisoalangiside (78),respectively. Thus, unlike monoterpenoid indole alkaloid biosyn-thesis, in A. lamarckii, the first enzymic condensation yields eitherthe (1R)- or (1S)-product, and naturally occurring (S)- and (R)-forms of the tetrahydroisoquinoline-monoterpene alkaloids andrelated nitrogenous glucosides.

1.14. Tryptophan-derived alkaloids

1.14.1. Simple indole alkaloidsPreviously contradictory results regarding the role of L-trypto-

phan in the biosynthesis of the indigo precursors, indican (79)


and isatan B (80), in four higher plants Baphicacanthus cusia (Nees)Bremek. (Acanthaceae), Calanthe veratrifolia R.Br. (Orchidaceae),Isatis tinctoria L. (Brassicaceae), and Polygonum tinctorium Alt.(Polygonaceae) were resolved by Xia and Zenk (1992) using[3-13C]indole and L-[3-13C]tryptophan. Indole and not L-tryptophanwas a biosynthetic precursor for the indoxyl derivatives.

The biosynthesis of the alkaloids gramine (1) and 2,4-dihy-droxy-2H-1,4-benzoxazin-3(4H)-one (DIBOA) (81) is mutuallyexclusive in Hordeum species (Poaceae). The cytochrome P450genes Bx2-Bx5 for DIBOA (81) biosynthesis in H. lechleri were func-tionally characterized and demonstrated a monophyletic origin forH. lechleri (Gruen et al., 2005). Homologous genes were found inother Hordeum species producing DIBOA (81), but not in speciesproducing gramine (1).

1.14.2. StrictosidineA number of studies on various aspects of the role of strictosi-

dine (82) in monoterpene alkaloid biosynthesis have been reportedin Phytochemistry over the years. Strictosidine (82) from Rhazyastricta Decsne. (Apocynaceae) was first described as the key inter-mediate in indole alkaloid biosynthesis in 1968, and preliminarybiosynthetic experiments were conducted by G.N. Smith (Univer-sity of Manchester) at that time, but not published. Based onpersonal witness the first publication to affirm that strictosidine(82), and not its 3b-isomer vincoside, was a precursor of tetrahy-droalstonine, ajmalicine (83), catharanthine (84), and vindoline(85) in Catharanthus roseus (L.) G.Don (Apocynaceae) was not re-ported until later by Brown et al. (1978).

Also in 1978, following the development of a radioimmunoas-say for ajmalicine (83), a soluble enzyme system from C. roseus cellsuspension cultures which synthesizes ajmalicine (83), 19-epi-ajmalicine, and tetrahydroalstonine from tryptamine and secolog-anin, was found to function optimally at pH 6.5. Alkaloid biosyn-thesis was inhibited by c-D-gluconolactone, a b-glucosidaseinhibitor, indicating the involvement of a b-glucosidase in theirbiosynthesis (Treimer and Zenk, 1978). The enzyme system wasnot substrate specific, and also utilized ring-substituted trypta-mines to catalyze the formation of unnatural ajmalicinederivatives.


In 1988, Hampp and Zenk (1988) reported on the first purifica-tion of strictosidine synthase, the primordial enzyme responsiblefor the condensation of secologanin and tryptamine to exclusivelyform 3a(S)-strictosidine (82). Using suspension cell cultures ofRauvolfia serpentina Benth. ex Kurz (Apocynaceae), the enzymewas purified 920-fold and had a specific activity of 184 nkat/mg.The enzyme was a single polypeptide, Mr 30,000, with a pH opti-mum of 6.5 and a temperature optimum of 45 �C. Apparent Km val-ues for both tryptamine and secologanin were 4 mM. Theimmobilized enzyme had a half-life of 100 days at 37 �C.

Verpoorte and co-workers (Stevens et al., 1993) purified tohomogeneity four isoforms of strictosidine synthase from a sus-pension culture of Cinchona robusta Howard (Rubiaceae). Themolecular weights were 35,000, 33,000, 35,000, and 33,000 dal-tons, and the pI values were 6.5, 6.5, 7.5, and 7.5, respectively.The Km values for the four enzymes for tryptamine were in themicromolar range. Inhibition of the enzymes was induced bystrictosidine (82), and by the subsequently formed quinoline alka-loids. The Ki of quinine for one of the main isoforms was 7 lM. Incontrast with strictosidine synthase from C. roseus, all four of theisoforms from C. robusta also accepted 5-methoxytryptamine as asubstrate.

Verpoorte and co-workers (Schripsema et al., 1991) administereda cell suspension culture from Tabernaemontana divaricata R.Br. exRoem. & Schult. (Apocynaceae) with 15N-labeled ammonium or ni-trate ion as alternative nitrogen sources. Ammonium ion was usedmore extensively than nitrate in the biosynthesis of the monoter-pene indole alkaloids. Considerably lower labeling percentages werefound for tryptamine than for O-acetylvallesamine (86) and the ami-no acids, suggesting the formation of a vacuolar pool of tryptamine.

Two cell lines of T. divaricata derived from the same suspension cul-ture were compared with respect to their biotransformation capac-ity (Dagnino et al., 1994). One of the cell lines accumulated indolealkaloids, mainly O-acetylvallesamine (86), at a high level, whereasthe other gave only low yields. When cultured with skeletally di-verse isolated alkaloids, such as conopharyngine, coronaridine,vobasine, or tabersonine, the alkaloids were similarly transformedby both cultures, indicating the same biotransformation potential.

[Me-2H3]Ajmalicine (83) degradation was investigated in a lowalkaloid-accumulating C. roseus cell culture, in comparison withchemical degradation in medium without cells. Degradation wasfaster in the presence of cells, being high at the beginning of thegrowth phase and in the stationary phase (dos Santos et al., 1994).

The second enzyme in the monoterpenoid indole alkaloid bio-synthetic pathway is strictosidine b-D-glucosidase (SG); and thisenzyme was partially purified from suspension cultured cells ofT. divaricata (Luijendijk et al., 1996). Three different forms werefound, each of which specifically catalyzed the deglucosylation ofstrictosidine (Km 0.2–1 mM). Some of the characteristics of the en-zyme were similar to those of the C. roseus SG enzyme. Cathen-amine (87), 21-hydroxyajmalicine (88), and isovallesiachotaminewere formed following 3a(S)-strictosidine (82) hydrolysis.

Biotransformation of 3a(S)-strictosidine (82), produced by het-erologously expressed strictosidine synthase from R. serpentina,with twenty-two bacterial strains demonstrated deglucosylationand rearrangement to vallesiachotamine (Shen et al., 1998). Thestudy indicates the potential for the biotechnological productionof alkaloids.

Camptothecin (CPT) (89) is a pharmaceutically importantmonoterpene indole alkaloid derived from strictosidine (82)


through a series of to-be-defined steps. A hairy root culture of Oph-iorrhiza pumila Champ. ex Benth. (Rubiaceae) produced highamounts of camptothecin (89), and it was shown that the site ofbiosynthesis is the endoplasmic reticulum. CPT is then transportedand localized in the vacuole, or, if the vacuole is blocked, CPT is ex-creted into the medium (Sirikantaramas et al., 2007).

Compared with other monoterpene indole alkaloids, relativelyfew studies have been reported on the alkaloids of Cinchona spe-cies. Levels of two of the early enzymes in the pathway, tryptophandecarboxylase (TDC) and strictosidine synthase (SS) were com-pared with the production of quinoline alkaloids in Cinchona ledge-riana Moens ex Trimen (Rubiaceae) suspension culturestransformed with A. tumefaciens (Skinner et al., 1987). Dark cul-tures showed higher levels of alkaloid production than light cul-tures. Stable levels of SS were up to 80 times higher wereobserved. The TDC activity declined after 13 days, and was consid-ered potentially rate limiting for alkaloid production.

It is often assumed that alkaloid biosynthesis is a consistent andevolving process from seed to seed. Verpoorte and co-workersshowed that for Cinchona alkaloids that is not the case (Aertset al., 1991). Germination of C. ledgeriana seeds produces a sub-stantial increase in alkaloid levels which only lasts about 4 days.Although the initial substrates, tryptophan, secologanin, and strict-osidine (82) are still all present, intracellular compartmentaliza-tion occurs which limits alkaloid biosynthesis. The concentrationof alkaloids present in the seedlings though was adequate to actas a feeding deterrent to slugs.

The history of strictosidine (82), and more particularly the dis-covery of strictosidine synthase as the key enzyme which catalyzesthe stereospecific formation of 3a(S)-strictosidine (82) from trypt-amine and secologanin is discussed in detail in an excellent reviewby Kutchan (1993). Aspects of the isolation and stereochemicaldetermination of strictosidine (82), the biochemical characteristicsof the enzyme, and how subsequent research led to new geneticapproaches to the study of plant enzymes and their expression insecondary organisms are presented. More than any other paperin the past 50 years in Phytochemistry, this review inherently de-picts the tremendous progress in the level of understanding ofalkaloid biosynthesis in plants.

The levels of five enzymes important in monoterpene indolealkaloid biosynthesis, isopentenyl pyrophosphate isomerase, gera-niol 10-hydroxylase, tryptophan decarboxylase, strictosidine syn-thase, and strictosidine glucosidase, were compared in two celllines of Tabernaemontana divaricate (Dagnino et al., 1995). OnlyG10H and SG showed a correlation with alkaloid biosynthesisand were considered rate-limiting. Added loganin stimulated thelow-alkaloid producing cell line 100-fold, indicating the depen-dence of alkaloid production on available monoterpene precursor.

1.14.3. AjmalineAlthough Rauvolfia serpentina Benth. ex Kurz (Apocynaceae)

plants accumulate ajmaline as the main monoterpene indole alka-loid, cell suspension cultures accumulate up to 1.6 g/l of the glu-coalkaloid raucaffricine (90), because the cell cultures do notcontain the enzyme raucaffricine-O-b-glucosidase (RG). Molecularcloning and functional expression of this enzyme in E. coli showedup to 60% amino acid identity with other glucosidases. Raucaffri-cine (90) was the best substrate for recombinant RG, and strictos-idine (82) was also a substrate (Warzecha et al., 2000).

1.14.4. Vindoline and related alkaloidsVindoline (85) is a major monoterpene indole alkaloid in the

leaves of C. roseus, and an important component of the anticancerbisindole alkaloids, vinblastine and vincristine (with the N-methyloxidized). A specific radioimmunoassay for vindoline (85) was

developed with a detection limit of 5 ng of the alkaloid (Kutneyet al., 1980b).

A multiple enzyme complex, involving at least five isozymes,which catalyzed the coupling of vindoline (85) and the iboga alka-loid catharanthine (84) to afford the bisindole alkaloid 30,40-anhy-drovinblastine (91) was partially purified from cell suspensioncultures of C. roseus (Endo et al., 1988).

Kutney and co-workers (1980a) examined the formation ofmonoterpene indole alkaloids in several cell lines of C. roseus.The 943 cell line produced a range of alkaloids, including vallesia-chotamine, isositsirikine, ajmalicine (83), yohimbine, hoerham-merinine, hoerhammericine, vindolinine, 19-epi-vindolinine, andthree new alkaloids, including two new tabersonine derivatives,19-acetoxy-11-methoxy-, and 19-hydroxy-11-methoxy-. No cath-aranthine (84) or vindoline (85) was detected.

Cryotechniques coupled with immunogold labeling was used tolocalize the intracellular storage of vindoline (85) in isolated pro-toplasts derived from the leaves of C. roseus. The label was presentin small vesicles and the cytoplasmic area bordering the plasma-lemma, as well as in the central vacuole and, to a much lesser ex-tent, in the chloroplasts (Brisson et al., 1992). These data supportthe difficulties encountered in the localization of vindoline (85)in leaf tissues.

S-Adenosyl-L-methionine:16-methoxy-2,3-dihydro-3-hydroxy-tabersonine N-methyltransferase (NMT) which catalyzes a latestep in vindoline (85) biosynthesis is localized in chloroplast thy-lakoids of C. roseus. The solubilized NMT was purified 120-foldand showed an apparent Mr of 60,000. However, it showed alteredspecificity for substrates compared with the membrane-associatedenzyme, and attempts at further purification resulted in loss ofactivity (Dethier and De Luca, 1993).

Metabolic profiling of 50 flowering cultivars of the ornamentalperiwinkle, C. roseus, by HPLC revealed similar levels of the mainmonoterpene indole alkaloids, with one cultivar showing a lowvindoline (85) content, which was traced to low levels of the en-zyme tabersonine 16b-hydroxylase (Magnotta et al., 2006).

1.14.5. Purine-derived alkaloidsThe biosynthesis of the common purine alkaloids, theophylline

(87), theobromine (88), and caffeine (89) in plants has remained asubject of significant investigation for many years. In one of theearly studies reported in the journal, leaf disks of Coffea arabica L.(Rubiaceae) were infiltrated simultaneously with L-methionine-methyl-14C, and with various possible precursors of caffeine bio-synthesis. The results permitted the identification of theobromine(88), 7-methylxanthine (90), and 7-methylxanthosine (91) as pre-cursors of caffeine (89) (Looser et al., 1974). 7-Methylguanosineseemed not to be an intermediate in caffeine (89) formation.

[8-14C]Adenine (92), rather than [8-14C]guanine (93), was amore effective precursor of caffeine (89) in Camellia sinensis (L.)Kuntze (Theaceae). Pulse labeling experiments with [8-14C]adenine(92) established that labeling of 7-methylxanthine (90) and theo-bromine (88) increased in the first 10 h, and then declined as caf-feine (89) labeling steadily increased, suggesting a possibleprecursor relationship (Suzuki and Takahashi, 1976).

Up to 26% of ring-labeled 7-methylxanthosine-14C (91) wasconverted into caffeine (89) after injection into leaves of C. arabicaover 8 days (Baumann et al., 1978). A possible caffeine biosyntheticpathway was discussed.

Cell-free extracts, with pH optima of 8.5, from partially ripe andunripe C. arabica fruits catalyzed the transfer of a methyl groupfrom S-adenosyl-L-methionine to 7-methylxanthine (90) and 7-methylxanthosine (91), producing theobromine (88). The samesystem also transferred a methyl group to theobromine (88) pro-ducing caffeine (89); Km values were determined (Roberts andWaller, 1979). The biogenetic pathway of caffeine formation in C.


arabica was proposed to be 7-methylxanthosine (91) ? 7-methyl-xanthine (90) ? theobromine (88) ? caffeine (89).

Cultured cells of C. sinensis could produce caffeine (89) andtheobromine (88), but at lower levels than cultured coffee cells(Furuya et al., 1990). [8-14C]Adenine (92) was used to show thatthe biosynthetic pathway to caffeine (89) was operative in the sta-mens and petals of C. sinensis, and that the reaction from theobro-mine (88) to caffeine (89) was present in C. sinensis and deficient inC. irrawadiensis Barua (Fujimori and Ashihara 1990). Purine alka-loid biosynthesis was not detectable in C. japonica L. and C. sasan-qua Thunb.

[8-14C]Adenine (92) was also used in field-grown leaf disks toexamine the annual seasonal variations in the metabolism of ade-nine nucleotides in the shoots of tea plants (Camellia sinensis cv.Yabukita) (Fujimori et al. 1991). Incorporation into theobromine(88) and caffeine (89) was found only in the young leaves. A leafextract catalyzed the methylation of xanthosine (94), 7-methyl-xanthine (90), and theobromine (88), but could not be detectedin extracts from leaves in which the synthesis of caffeine (89)was not observed.

Caffeine (89), theobromine (88), and theophylline (87) levels inaqueous extracts of the endosperm of the immature and maturefruits of C. arabica and six other species of Coffea were examinedby HPLC (Mazzafera et al., 1991). Caffeine (89) was the dominantalkaloid, and the highest levels were found in C. canephora Pierreex. A.Froehner (Rubiaceae) at 35.1 and 24.5 mg/g, respectively, inthe immature and mature endosperm. Caffeine (89) was not de-tected in the mature fruits of C. bengalensis Roxb. The rate of turn-over of caffeine (89) in various species of Coffea was examinedusing [8-3H]caffeine (89). It was metabolized slowly by the imma-ture endosperm of C. arabica and C. canephora, and more rapidly byC. dewevrei De Wild. & T.Durand., C. eugenioides S.Moore, C. steno-phylla G.Don, C. salvatrix Wynn. & Philipson, and C. benghalensisB.Heyne ex. Schult. Potential sources for a natural decaffeinatedcoffee were discussed.

Waller and co-workers provided an authoritative review of themetabolism of the purine alkaloids caffeine (89) and theobromine(88) in coffee and tea plants (Suzuki et al., 1992). The activity of thethree N-methyltransferases was discussed, together with the dif-ferent rates of metabolism in the various species of the two genera

at different stages of growth. The physiological role of the purinealkaloids in the plant remains elusive,

[8-14C]Adenine (92), [9-14C]guanosine (95), and [8-14C]hypo-xanthine (96) were incorporated into theobromine (88) and caf-feine (89) in the young, but not the old, leaves of mate [Ilexparaguariensis A.St.-Hil. (Aquifoliaceae)] (Ashihara, 1993). The con-ventional pathway for the degradation of purines via allantoin andallantoic acid was found to operate.

Caffeine (89) and theobromine (88) were trace components, andtheophylline (87) was absent, when the seeds and leaves of nineHerrania Goudot (Sterculiaceae) and eleven Theobroma L. (Sterculi-aceae) species were examined. Tetramethyl urate (97) was theprincipal purine alkaloid detected (Hammerstone et al., 1994).

Conversion of [8-14C]adenine (92) to theobromine (88) and caf-feine (89) was found only in the first leaves of coffee plants, no the-ophylline (87) was detected (Fujimori and Ashihara, 1994).Secondary leaves produced nucleotides and nucleic acids.

The purification and the N-terminal sequence of S-adenosyl-L-methionine: theobromine 1-N-methyltransferase (STM), the en-zyme responsible for the methylation of theobromine (88) to caf-feine (89) in coffee was described (Mazzafera et al., 1994). Theenzyme showed an apparent Mr of ca. 54,000 and ca. 60,000 bygel filtration and SDS–PAGE, respectively. Using theobromine(88) as substrate, the Km value for S-adenosyl-L-methionine was10 lM. It is a bifunctional enzyme which also methylates 7-meth-ylxanthine (90), the precursor of theobromine (88). The N-terminalsequence for the first 20 amino acids was obtained for STM, but nosimilarities were found with other methyltransferases.

Significant challenges were encountered in demonstrating theconversion of xanthosine (94) to 7-methylxanthosine (91) in sus-pension cultures of coffee. Cells administered with 5 mM ethephonor 1 mM adenine to enhance purine alkaloid formation increasedthe incorporation of [14C]adenine (92) into the purine alkaloids,as did ethephon (Schulthess and Baumann, 1995a). Neither ethe-phon nor adenine treatment led to a distinct incorporation of[14C]adenine (92) or [methyl-14C]methionine into 7-methylxan-thosine (91). Kinetic analysis based on [14C]adenine (92) indicatedthat xanthosine (94) was not a precursor in caffeine (89) biosyn-thesis. It was postulated that metabolic channeling occurred inpurine alkaloid biosynthesis, including the formation of xanthosine


(94), its methylation to 7-methylxanthosine (91), and finally enzy-matic hydrolysis to 7-methylxanthine (90).

In the dark, both adenine (92) and ethephon stimulated purinealkaloid [7-methylxanthine (90), theobromine (88), and caffeine(89)] formation by factors of 4 and 7, respectively in cell suspen-sion-cultures of coffee as determined by HPLC. In light, caffeine(89) formation was enhanced by a factor of 21 without affectingtheobromine (88) and 7-methylxanthine (90) pools. Co-feedingadenine feeding resulted in enlarged nucleotide pools (xanthosine,guanosine, and inosine) and 7-glucopyranosyladenine (Schulthessand Baumann, 1995b).

Although young coffee leaflets were actively producing (about38 or 71% of the recovered radioactivity from [U-14C]adenine(92) and [Me-14C]methionine, respectively), 7-methylxanthine(90), theobromine (88), and caffeine (89), no incorporation into7-methylxanthosine (91) or a monomethylated purine other than7-methylxanthine (90), was observed (Schulthess et al., 1996).The result was similar to that obtained with coffee cell suspensioncultures, implying that the first methylation step in caffeine (89)synthesis is metabolically channeled. When the methyltransferaseactivity was examined, only xanthosine (94) and xanthosine 50-mono-phosphate (XMP) (98) were methylated, and exclusively atthe N-7 position. The new XMP N-methyltransferase activity wasmeasured under conditions stabilizing the product. As a result, anew hypothesis regarding caffeine (89) biosynthesis which favorsmethylation of XMP (98) rather than xanthosine (94) as the crucialstep which leads from primary metabolism into caffeine (89) syn-thesis was proposed.

The d13C- and d15N-values of caffeine (89) and theobromine (88)permit distinction between natural and synthetic products. The 13Cpattern of the natural purine alkaloids indicated an enrichment of13C in the positions 2 and 8, originating from the THF pool and adepletion in the methyl groups derived from the SAM pool (Weil-acher et al., 1996). These results suggest the presence of two sep-arate C1 pools.

The last two steps in caffeine (89) biosynthesis in coffee are theconversion of 7-methylxanthine (90) to theobromine (88), andsubsequently of theobromine (88) to caffeine (89) by two N-meth-yltransferases. Waldhauser et al. (1997a) devised an assay for theseenzyme activities using high concentrations of methyl donor andsubstrate, obviating the need for tracer studies. The enzymesshowed maximum stability at pH 7.5, and at around 35 �C. Maxi-mum relative and absolute methyltransferase activities coincidedwith leaf emergence, when, as expected, chemical defense hasthe highest priority and the purine alkaloids are at peak

concentrations. The transient accumulation of theobromine (88)in the young leaflets supported the observed competition of thetwo NMT activities for SAM.

The first N-methylation step in the caffeine biosynthetic path-way is the formation of 7-methylxanthine (90). Using young,emerging coffee leaflets, anion exchange chromatography andchromatofocusing separated XMP N-7-methyltransferase from 7-methylxanthine N-3-methyltransferase, and theobromine N-1-methyltransferase (Waldhauser et al. 1997b). The enzymes wereseparated by photoaffinity labeling with [methyl-3H]SAM, fol-lowed by SDS–PAGE, and eventually provided all three N-methyl-transferase activities in bands at 49, 43, and 40 kDa. Correlationsof the 49- and 43-kDa bands with the 2nd and 3rd NMT activitieswere not established.

Caffeine (89) and theobromine (88) accumulated in callus androot suspension cultures in MS medium from C. sinensis as estab-lished by TLC, UV, and GC (Shervington et al., 1998). In prepara-tions from young leaves of C. sinensis, the N-3-methyltransferasewas associated with a purified chloroplast preparation (Katoet al., 1998).

Caffeine (89) biosynthesis from [8-14C]adenine (92), and itsoverall metabolism were investigated in anti-sense and RNA inter-ference transgenic plants of C. canephora in which the expression ofCaMXMT1 was suppressed. Synthesis from adenine (92) and con-version of theobromine (88) to caffeine (89) were both reducedin the transgenic plants, and metabolism of [8-14C]adenine (92)shifted from purine alkaloid synthesis to purine catabolism or sal-vage for nucleotides (Ashihara et al., 2006).

Ashihara et al. (2008) provided a review which details how thebiosynthetic pathway of caffeine involves the conversion of xan-thosine (94) ? 7-methylxanthosine (91) ? 7-methylxanthine(90) ? theobromine (88) ? caffeine (89). The genes encoding N-methyltransferases involved in three of these four reactions havebeen isolated and their molecular structures investigated. Path-ways for the catabolism of caffeine (89) have thus far not producedenzymatic or genetic studies. The metabolism of purine alkaloidsin Camellia, Coffea, Theobroma, and Ilex plants was summarized,and the evidence for the involvement of caffeine (89) in chemicaldefense and allelopathy presented. Studies on metabolic engineer-ing to produce coffee seedlings with reduced caffeine (89) content,and transgenic caffeine-producing tobacco plants with enhanceddisease resistance were also discussed.

1.15. Miscellaneous alkaloids

1.15.1. CyclopeninIn 1976, the enzyme preparation cyclopenin m-hydroxylase was

obtained from Penicillium cyclopium and transformed cyclopenin(99) into the hydroxy derivative cyclopenol (100) (Richter and Luck-ner, 1976). The enzyme required molecular oxygen and NADPH foractivity, and was inhibited by both thiocyanide and cyanide ion.

Three of the enzymes, cyclopeptine dehydrogenase, dehydrocy-clopeptine epoxidase, and anthranilate adenyltransferase, in cylco-penin (99) biosynthesis were induced endogenously in the hyphaeof P. cyclopium leading to 10-fold increase in alkaloid production(Voigt et al., 1978).

1.15.2. Holaphyllamine[4-14C]Holaphyllamine (101) was converted to pregnenolone

(102) by the leaves of Holarrhena floribunda T.Durand & Schinz(Apocynaceae), and completed a cycle in which pregnenolone(102) had been converted to holaphyllamine (101) (Bennettet al., 1966).


1.15.3. Ricinine and HermidinOne of the earliest plant alkaloid enzyme preparations was

developed by Robinson (1965) from Ricinus communis L. (Euphorbi-aceae) which converted 1-methylnicotinonitrile (103) to the corre-sponding 4- and 6-pyridones.

The biosynthesis of the pyridinone hermidin (104) was studiedthrough [U-13C6]glucose and 13CO2 pulse labeling experiments inMercurialis annua L. (Euphorbiaceae) (Ostrozhenkova et al., 2007).Based on similar labeling profiles, the results supported the deriva-tion of hermidin (104) through the nicotinate pathway, by way ofdihydroxyacetone phosphate and aspartate. The pulse/chase appli-cation of 13CO2 permits experiments under physiological condi-tions and should be widely applicable.

2. Conclusions

An overview of the studies published in Phytochemistry in thepast 50 years related to the biosynthesis of alkaloids has been pre-sented. Techniques for the study of alkaloid biosynthesis haveevolved dramatically in this period from studies of the utilizationof potential precursors to radio- and then stable-isotope studies,to the isolation of crude and then purified enzyme systems. Nowstudies are routinely providing important information regardingthe functionality of the genes which are responsible for the en-zymes of alkaloid biosynthetic pathways. Progress on several ma-jor alkaloid biosynthetic pathways has appeared inPhytochemistry in the past 50 years, and the journal has thereforeplayed a crucial role in advancing the science and the technologybehind the determination of how nature produces alkaloids inplants.

Acknowledgments

The author acknowledges many alkaloid teachers who in thepast 45 years have provided knowledge and inspiration regardingthe biosynthesis of these amazing natural products which are suchan integral aspect of all our lives. The author also acknowledges theassistance of Dr. Nabil Ali Al-Mekhlafi, Institute of Bioscience, Uni-versity of Putra Malaysia, Selangor, Malaysia in the preparation ofthe diagrams for this article.

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Professor Emeritus Geoffrey A. Cordell obtained a Ph.D.in synthetic natural product chemistry at the Universityof Manchester. After joining the College of Pharmacy atUniversity of Illinois at Chicago he directed drug dis-covery programs, served as a Department Head andInterim Dean, and held several senior research admin-istrative positions at the College and Campus levels; heretired in 2007.He has about 600 publications, is the editor of 37 books,including 29 volumes of ‘‘The Alkaloids: Chemistry andBiology’’, and has lectured extensively around the world.He is a member of the Editorial Advisory Board of

twenty-five scientific journals, is one of 14 Honorary Members of the AmericanSociety of Pharmacognosy, and is an Honorary Professor at Sichuan University,Chengdu, PRC.
After over 45 years studying the isolation, structure elucidation synthesis, andbiosynthesis of natural products, his efforts are now focused on developing naturalproduct research programs, where he serves as an advisor and consultant in severalcountries in various parts of the world. His interests include strategies for thesustainability and quality control of medicinal agents, the detection of biologicallyactive natural products, the chemistry and biosynthesis of alkaloids, and the use ofvegetables as chemical reagents.

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Fifty years of alkaloid biosynthesis in Phytochemistry · 2014. 8. 13. · Fifty years of alkaloid biosynthesis in Phytochemistryq Geoffrey A. Cordell⇑ Natural Products Inc., Evanston,